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Article

Pine Bark as a Potential Source of Condensed Tannin: Analysis through Fourier Transform Infrared Spectroscopy (FTIR), Scanning Electron Microscopy (SEM), and Energy Dispersive X-ray (EDX)

by
Rossy Feria-Reyes
1,
Sergio Obed Ramírez-Cruz
2,
Faustino Ruiz-Aquino
2,*,
Luis Humberto Robledo-Taboada
3,
Marco Antonio Sánchez-Medina
4,
Oscar Francisco Mijangos-Ricárdez
2,
Rosalío Gabriel-Parra
2,
Mario Ernesto Suárez-Mota
2,
Ramiro Puc-Kauil
5 and
Jhazeel Porcallo-Vargas
6
1
Department of Electronic Engineering, Tecnológico Nacional de México/Instituto Tecnológico de Oaxaca, Calz. Tecnológico No. 125, Oaxaca C.P. 68030, Mexico
2
Instituto de Estudios Ambientales, Universidad de la Sierra Juárez, Avenida Universidad S/N, Ixtlán de Juárez, Oaxaca C.P. 68725, Mexico
3
Department of Industrial Engineering, Tecnológico Nacional de México/Instituto Tecnológico de Oaxaca, Calz. Tecnológico No. 125, Oaxaca C.P. 68030, Mexico
4
Department of Chemical and Biochemical Engineering, Tecnológico Nacional de México/Instituto Tecnológico de Oaxaca, Calz. Tecnológico No. 125, Oaxaca C.P. 68030, Mexico
5
División de Ingeniería Forestal, Tecnológico Nacional de México/Instituto Tecnológico Superior de Venustiano Carranza, Av. Tecnológico S/N, Colonia El Huasteco, Puebla C.P. 73049, Mexico
6
Facultad de Medicina y Cirugía, Universidad Autónoma Benito Juárez de Oaxaca, Oaxaca de Juárez, Oaxaca C.P. 68120, Mexico
*
Author to whom correspondence should be addressed.
Forests 2023, 14(7), 1433; https://doi.org/10.3390/f14071433
Submission received: 18 June 2023 / Revised: 2 July 2023 / Accepted: 7 July 2023 / Published: 12 July 2023
(This article belongs to the Section Wood Science and Forest Products)
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Abstract

:
This study aimed to evaluate the tannin content in the bark of five pine species from a forest harvesting area of the Ixtlán de Juárez community, Oaxaca, México. The species studied were Pinus patula, Pinus ayacahuite, Pinus rudis, Pinus douglasiana, Pinus pseudostrobus. The bark samples were subjected to a drying, grinding, and sieving process. These compounds were extracted using two methods: by constant reflux with ethanol for three hours or in a water bath with distilled water for two hours. The percentage of total extract, Stiasny number, and condensed tannins were quantified. The quantitative number of condensed tannins present in the bark for each of the five species studied ranged from 0.65% to 5.14% for the ethanolic extracts and 0.14% to 1.46% for the aqueous extracts. Analysis by Fourier transform infrared spectroscopy (FTIR) identified polyphenolic compounds and functional groups characteristic of tannins. Analysis by scanning electron microscopy (SEM) and X-ray dispersive spectroscopy (EDX) provided the elemental analysis and chemical composition of the tannin extracts, respectively, finding trace elements of silver, cadmium, phosphorus, iodine, and sulfur, which are minerals present in the soil, meaning that through processes of micronutrient absorption, these can interact with the tannins or polyphenols in the barks of the tree species being studied. The results indicate that the bark of P. patula and P. ayacahuite have a higher concentration of condensed tannins, 5.14% and 4.71%, respectively, in the ethanol extraction, and may be susceptible to exploitation due to the amount of bark generated in forestry activities.

1. Introduction

The Sierra Norte region in the state of Oaxaca is one of the most important forest-producing regions in Mexico. This region has an extension of 379, 034.75 hectares of forest area [1]. Its forests are located at altitudes of between 2500 and 3000 m above sea level (masl), where pure forests of Pinus rudis Endl., P. patula Schiede ex Schltdl. & Cham., P. ayacahuite Enhrenb ex Schltdl., P. pseudostrobus Lindl., P. oaxacana Mirov can be found, alongside forests with mixtures of these pine species and oak of various species, in addition to mountain mesophilic forests [1]. The community of Ixtlan de Juarez, Oaxaca, has an important timber potential due to its 19,280.14 ha of forest area [2] and the high productivity of its forest [3].
Forest residues, such as sawdust and tree bark, are generated as a result of forest harvesting [4]. Bark corresponds to a group of non-timber forest products that are widely used and underutilized [5].
Tannins are present in all plant cells and are found mainly in the soft tissues of plants, such as the leaves, needles, and bark of different tree species, including pine (pinus sp.), oak (Quercus sp.), and other tree species, such as Arbutus xalapensis and Prunus serotina [6,7].
The bark is the second most important tissue in the trunk after the wood, potentially representing about 10%–20%, depending on the species and growing conditions [8,9]. Similar to wood, the bark is chemically composed of typical cell wall components: cellulose, lignin, and extractable substances formed from the secondary metabolism of plants [10]. However, the chemical composition of bark is more complex than wood composition, which is mainly distinguished by the high content of polyphenols and suberin. Polyphenols include simple phenols, such as lignans, stilbenes, flavonoids, quinones, and tannins [11]. Figure 1 shows a classification of tannins, including floranthanins, a type of tannin found in algae in the Phaeophyta phylum, such as sargassum. In contrast to hydrolysable or condensed tannins, these compounds are oligomers of phloroglucinol [12]. Phlorotannins are classified into fuhalols, floretols, fucols, fucofloretols, echols, and carmalols.
Tannins are polymeric phenolic compounds that are produced in plants as metabolites and have the ability to form complexes with proteins, polysaccharides, nucleic acids, steroids, and saponins [9]. The type and content of the tannins vary due to various factors being analyzed, such as age, species, tree conditions, location, and the section or area of the tree [13].
Tannins have various applications. They are used in manufacturing chemical products, inks, and pharmaceuticals, alongside as a clarifying agent in winemaking. Moreover, they are used in medicine as an antidiarrheal, in healing, and in other applications [13]. They are effective for tanning animal hides in the leather processing industry [14], mineral absorption [15,16], and protein precipitation purposes [17]. Furthermore, they are also used for producing adhesives in the woodwork industry and anticorrosion chemicals [18].
The main industrial use of tannins is leather tanning, while the second most important application of vegetable tannins is in the formulation of wood adhesives [19], whereby at the industrial level, the different technologies require energy for tannin production operations from plant species, optimizing their operating parameters with minimum energy consumption, mainly in the operations of extraction, concentration, distillation, and drying of the product [20].
In the extraction of tannins, various solvents, and mixtures have been used at various temperatures, such as acetone–water [21], acetone–ethanol [22], methanol–water [23], and methanol–water using an ultrasound system [24], water–sodium sulfite [25], water–NaOH [26], and water [27,28]. In general, tannins tend to dissolve better in polar solvents. However, the use of these solvents turns out to be highly expensive and can sometimes be toxic. Thus, extraction techniques must be more innovative and have a low environmental impact, such as shorter extraction times and a minimum number of solvents, which leads to lower energy consumption and economic costs [29].
In addition to its traditional uses, pine bark can have multiple applications that can increase the bark value chain due to the considerable presence of extractable substances in its tissue. It has been identified that softwood bark extracts are rich in phenolic compounds, especially in condensed tannins. Therefore, the objective of the present work was to evaluate the tannin content through two extraction methods and characterize the extracts of five pine species (Pinus patula Schiede ex Schltdl. et Cham., Pinus ayacahuite Enhrenb ex Schltdl., Pinus rudis Endl, Pinus douglasiana Martínez, and Pinus pseudostrobus Lind.) from a forest harvesting area in the community of Ixtlan de Juarez, Oaxaca, Mexico, using infrared spectroscopy and scanning electron microscopy.

2. Materials and Methods

2.1. Study Area

The study was conducted in the forests of the Ixtlan de Juarez community, Oaxaca, Mexico. The community is located in the Sierra Norte region at geographic coordinates of 17°18′16″ N, 96°20′00″ W and 17°34′00″ N, 96°31′38″ W (Figure 2), at an altitude of 2300 masl [2]. It has a land area of 19,310.14 hectares, of which, 12,389.50 hectares are temperate forests, and 6890.64 hectares are mountain mesophilic forests and jungles [2]. The composition of their commercial forests is primarily pine–oak [30]. On the other hand, within the forests of the Ixtlan de Juárez district, Oaxaca, there are 23 species of the Quercus genus [31] and 13 species of Pinus [32].
The soil type in the forests was mostly Luvisol (41.35%), Acrisol (35.62%), Cambisol (21.98%), and Fluvisol (1.05%) [33]. The temperature ranged from 10 to 26 °C, with a precipitation range of 700 to 4000 mm. According to the Köppen classification, modified by [34], the climate of the pine–oak forest was C (m) (w″) b (i′) g, humid temperate with summer rainfall.

2.2. Tree Selection and Sample Preparation

One tree of each species was chosen: Pinus patula Schiede ex Schltdl. et Cham., Pinus ayacahuite Enhrenb ex Schltdl., Pinus rudis Endl., Pinus douglasiana Martínez, and Pinus pseudostrobus Lind. The trees with the best phenotypic characteristics, such as straight stem, cylindrical stem, and superior growth were used. For each species, 5 kg of bark was collected and crushed in a Wiley® mill, then, homogenized using a 1.0 mm opening sieve. The moisture content of the samples was determined using a Riossa® drying oven at 70 °C for 24 h.
The dasometric data and geographical location of the trees are shown in Table 1.

2.3. Extraction of Tannins with Ethanol, Stiasny Number, and Condensed Tannins

For each species, 25 g of bark was separated and 200 mL of 95% ethanol was added. The mixture was refluxed in a Soxhlet system for 3 hours at 80 °C. The extractions were carried out in triplicate. Subsequently, the samples were hot vacuum-filtered in a Büchner funnel, using Whatman® No. 54 filter paper (22 μm pore size). The filtered extracts were concentrated in a RE100-Pro rotary evaporator and dried in a Riossa® drying oven at 50 °C for 12 h. The amount of material extracted was determined by the difference in weight before and after extraction of the sample,.
The Stiasny number was determined using the dried ethanolic extracts in triplicate, while 0.1 g of extract was weighed. Then, using a 200 mL Erlenmeyer flask, 10 mL of 99.8% methanol was added [7].
A total of 1 mL of concentrated HCl and 2 mL of 37% formaldehyde were added to the resulting mixture. Then, it was heated at 100 °C for 30 min. Next, the mixture was filtered under a vacuum in a Büchner funnel, using Whatman No. 54 filter paper. The obtained solid residue was dried at 105 °C until a constant mass was reached.
The mass of the solid residue corresponded to the amount of condensable tannin present in the dry extract. It is expressed as a percentage, with respect to the initial mass of the dry extract, as established in the expression of the Stiasny number Equation (1).
S t i a s n y   n u m b e r =   s o l i d   r e s i d u e   d r y   m a s s i n i t i a l   s a m p l e   d r y   m a s s   × 100
The condensed tannins were calculated quantitatively, according to Equation (2) [35]:
C T = S N × T E 100
where CT was condensed tannins, NS was the Stiasny number, and TE represented the total extract.

2.4. Extraction of Tannins with Distilled Water, Stiasny Number, and Condensed Tannins

The extractions were carried out in triplicate. A total of 5.0 g of bark was used from each sample. A total of 75.0 mL of distilled water and 100 μL of glacial acetic acid were added to a 250 mL Erlenmeyer flask immersed in a Riossa® water bath at 87 °C. Subsequently, the cold extract was filtered through a Büchner funnel using Whatman® No. 54 filter paper. The filtered solution was adjusted to 100 mL [36,37], as shown in Figure 3.
The total extract was calculated by gravimetry. Next, 50 mL of the filtered solution was placed in a 50 mL beaker and dried in the oven at 105 °C. The total extract was quantified as the remaining material.
Subsequently, 5.0 mL of formaldehyde and 2.5 mL of HCl were added to the remaining volume (50 mL) of the filtered solution for over 30 min using a Soxhlet system. Then, the mixture was vacuum filtered through a Büchner funnel, using Whatman® No. 54 filter paper, and dried at 105 °C for 24 h.
The Stiasny number is the ratio between the precipitate formed with respect to the total solids and corresponds to the percentage of condensed tannins in the extract [35]. It was calculated using Equation 1. The percentage of condensed tannins in the bark was obtained by multiplying the Stiasny number by the yield in the solids obtained in each extract, according to Equation (2).

2.5. Characterization of Tannins by Fourier Transform Infrared Spectroscopy (FTIR)

The extracts obtained from the bark of each species were characterized by infrared spectroscopy, using a PerkinElmer® Frontier™ FTIR spectrophotometer in a wave number range between 500 and 4500 cm−1 and via the diffuse reflectance method.

2.6. Analyses by Scanning Electron Microscopy (SEM) and Energy Dispersive X-ray (EDX) Spectrometry

For this analysis, a Thermo Fisher Scientific (Waltham, MA, USA) Phenom Pro-X scanning electron microscope was used with 10 mm diameter aluminum pins. A double-sided copper conductive appointment was placed on the pin, and the samples (in this case powders) were placed. The equipment has a 10 kV field emission cannon, which allows for the chemical characterization and/or elemental analysis of the materials. Therefore, the elemental analysis of each of the tannins of the tree species obtained was determined using the same equipment. Thus, the elemental and chemical characterization of the tannin samples obtained from the five tree species was carried out using X-ray energy dispersion spectroscopy.

2.7. Statistical Analysis

The data were analyzed using a completely randomized design. The quantitative variables were total extract, Stiasny number, and condensed tannins, with 15 repetitions per variable; the studied species were used as treatments.
An analysis of variance (ANOVA) was carried out using the GLM procedure to determine the significant differences between the extractive yield of the five studied species.
Subsequently, a Tukey test was carried out to compare the means obtained and determine the species that presents the best extractive yield. The statistical analyzes were carried out using the SAS statistical program [38] version 9.0 with a significance level of 95% (α = 0.05).

3. Results and Discussion

3.1. Tannin Extraction with Ethanol, Stiasny Number, and Condensed Tannins

Significant differences were obtained among the number of obtained extracts from the five studied species. The total extract values (Table 2) were within the interval delimited by the maximum (14.53%) and minimum (4.92%) values belonging to P. douglasiana and P. pseudostrobus, respectively. In some extraction processes, 100% ethanol is used as a solvent for the P. patula bark, obtaining 9.37% of the total extract, while for P. oocarpa it obtained 6.80% [39]. In this study, the P. douglasiana species shows an increase in the mentioned values, with 14.53% of the total extract. In other studies, they used a 50% ethanol–water mixture for the extraction of tannins from 8 pine species, reporting extractions of 9.66% for P. arizonica, 19.39% for P. ayacahuite, 9.94% for P. cooperi, 8.86% for P. chihuhuana, 16.68% for P. duranguensis, 6.66% for P. engelmannii, 19.11% for P. leiophylla, and 12.125% for P. teocote [35]. Similarly, for P. caribea, an average tannin value of 19.34% was reported in material extracted using 95% ethanol [11]. The variation in extraction yields depends on the species and factors such as particle size, bark-solvent ratio, agitation, extraction time, type of solvent used, and temperature [39].
In the present study, mature trees with an average age of 47 years were analyzed; however, the age of the tree plays an important role in obtaining the tannins [40]. In the bark of juvenile Acacia melanoxylon trees, a higher number of total extractives (29%) were extracted with solvents of increasing polarity (dichloromethane, ethanol, and water), followed by mature bark (21%), because juvenile tissues are more phytochemically active than mature tissues [40]. Moreover, alongside age, the relative volume and thickness of the bark decreases in the direction from the rump to the top [41]. Furthermore, it influences the quality of the tannins, the harvesting season (winter and summer), and the extraction procedure [42].
Similarly, polyphenol concentration is due to natural variability, such as genotype, differences in growth, climate, and soil type [43]. Poor soil quality increases the number of tannins in plants [44]. Soil nitrogen is negatively correlated to extractives and lignin, whereby, if the nitrogen level decreases in the soil, the extractives and lignin content increases in the wood [45]. Additionally, the soil factors influence tree development, with the macronutrients nitrogen and phosphorus having the highest demand in plants over time [46]. Soil nitrogen affects both anthocyanin and flavonoid content, and generally higher polyphenol content was observed when less nitrogen fertilizer was added to the soil [47].
Using the Stiasny number, significant differences were found between the analyzed species. P. patula (Table 2) presented the highest mean, while P. rudis presented the lowest at 68.14% and 8.61%, respectively.
These values are low compared to those obtained using a 50% ethanol–water mixture as a solvent for P. ayacahuite and P. cooperi, which presented Stiasny numbers of 80% and 71%, respectively [35]. Tannins presenting Stiasny numbers higher than 65% are considered a good source of adhesives in cellulosic or wood products applications [48]. Thus, the P. patula species, which presents a Stiasny number of 68.14%, fulfills this criterion. However, it is necessary to consider the number of condensed tannins presented by each species being studied to determine the optimal species that fulfills this function, since there are significant differences between each of them. The P. patula species presented a higher average number of condensed tannins (Table 2), with a value of 5.14%. Conversely, P. rudis presented the lowest average number of condensed tannins at 0.65%.
Some studies have reported that P. caribaea presents an average value of condensed tannins of 13.67% when extracted using 95% ethanol [49]. On the other hand, when extracting tannins from 8 pine species with 50% aqueous ethanol, the reported total extract values were from 6.46% to 16.18% [35]. The reported results were higher in both cases than for those obtained in the five species analyzed in this study. These low Stiasny number values can be attributed to extraction time and temperature, particle size, and type of bark, which promote variations in extraction yields and condensed tannin contents [39].

3.2. Extraction of Tannins with Water, Stiasny Number, and Condensed Tannins

Significant differences were presented in the Stiasny number in all cases when the extraction process used distilled water. The values varied from 1.16% to 3.19% for P. pseudostrobus and P. douglasiana, respectively (Table 3).
Total extracts from the thermally treated samples were between 4.04% and 4.97% when the extraction was conducted using distilled water. [50]. When hot water was used to extract tannins from the bark of 8 pine species, values were reported in the range of 2.23% to 10.61% [35].
The difference in values in the different tannin extracts in each species was lower in the aqueous extract than in the ethanolic extract due to the polarity of each solvent being used [50].
For the Stiasny numbers, significant differences were found among the analyzed species, whereby P. patula (Table 3) was the species with the highest average, with a value of 64.60%. On the other hand, the species with the lowest average Stiasny number was P. rudis, with a value of 8.18%.
These values are very similar to those reported by other researchers, where they found a Stiasny number of 69% for P. ayacahuite, P. cooperi with 44.4%, and Pinus engelmannii with 30%, using an extraction system where hot water was the solvent [24]. On the other hand, the extraction of tannins using distilled water from the bark of thermally treated pine species provided a Stiasny number of 78.29% [51]. Furthermore, for aqueous extracts of P. radiata bark, a Stiasny value of 77.20% was found [52]. The present study determined that P. patula (64.60%) is the species closest to the established criteria that can be considered an essential source of condensed tannins [49]. However, it is important to consider the amount of bark generated and provide it an added value by obtaining tannins. In the number of condensed tannins, there were significant differences between each of the five species, where P. patula presented a higher mean of condensed tannins (Table 3), with a value of 1.46%. On the contrary, P. rudis presented a lower mean of condensed tannins, with a value of 0.14%. Different studies have reported condensed tannin yields of 10.5% for an aqueous extract of P. radiata [53]. Furthermore, in the extraction of tannins from the bark of thermally treated pine species, they obtained a condensed tannin value of 3.43% after performing the extraction with distilled water at the best treatment heat [51].
Due to the polarity of each solvent used, it was notable that the condensed tannin content obtained in the ethanolic extracts was higher than in the aqueous extracts [35,51].
The percentages obtained in this study are relatively low. However, it should be taken into account that, when working with matter such as tree bark, it is necessary to consider not only the number of condensed tannins present in the lyophilized extracts but also the amount of material extracted from the bark, since this is the direct source of these metabolites [49].

3.3. Fourier Transform Infrared Spectroscopy (FTIR) Analysis

The tannins obtained from the five tree species were analyzed by Fourier transform infrared spectroscopy (FTIR). These tannin samples were compared using the HPLC grade catechin standard provided by Sigma Aldrich [37]. Each spectrogram of the tannin extracts obtained using ethanol was similar and presented a wide and strong band between 3220 and 3430 cm−1. This was attributed to the stretching vibrations of the hydroxyl groups -OH present in the phenolic compounds [54].
The aqueous extracts showed a similar band centered around 3270 cm−1 (Figure 4). Similar results were reported when the tannins were present in the bark of Pinus patula, where a moderate band was also presented between 3400 and 3200 cm−1 [55]. In addition, signals at 3421 and 3024 cm−1 were attributed to hydroxyl (-OH) groups in the analyzed spectra of cotton (Gossypium hirsutum L.) and wheat straw (Triticum aestivum L.) stems [37].
In the ethanolic extracts that were analyzed from the different tree species, a peak can be observed around 2918 cm−1 and another more moderate one at around 2850 cm−1, which can be associated with the symmetric stretching vibrations and antisymmetric -CH- of the CH2 and CH3 groups, as reported by [29].
Similarly, in the aqueous extracts, similar peaks were observed at around 2932 cm−1 and 2840 cm−1. Similar results were identified at 2900 cm−1, which were attributed to the stretching characteristics of C-H [36].
For the ethanolic and aqueous extracts, the signals around 1603 cm−1 and 1513 cm−1 were assigned to the deformation vibrations of the carbon–carbon bonds in the region of 1605–1496 cm−1, in the aromatic structures present in these types of metabolites [36].
Some signals were identified at 1270 cm−1, which correspond to the C-O bond tension as well as the bonds of the carboxylic groups, with very pronounced peaks at around 1040 cm−1, resulting from the vibrational stretching effect of the C–O bonds present in phenols [49,52]. Finally, we can identify moderate signals in both extracts around 660 cm−1 that corresponds to C-H bending vibrations [37].
When the tannin extraction was carried out using alcohol, additional reactions occurred in the polyphenols selectively. This implies that the vibrational waves present greater intensity in the various functional groups, with respect to the aqueous medium.
The C=O stretching frequencies of the aldehydes were similar to the ketones.
The C=H stretching of the CH=O group in aldehydes appeared as a pair of bands in the interval range of 2700–2900 cm−1 [56].
In the case of esters, there was a strong C=O absorption at 1730 and 1750 cm−1. Esters exhibit absorptions for symmetric and antisymmetric stretching of C-O-C in ranges from 1050 to 1300 cm−1.
However, ether functional groups are characterized by an intensity and broadband due to the C-O-C antisymmetric elongation and range from 1070 to 1150 cm−1.
Ethers of the ROR’ type, where R and R’ are different, have two regional absorptions. The most characteristic bands in the IR spectra of carboxylic acids are those of the hydroxyl and carbonyl groups and a broad signal for O-H stretching of 2500 to 3600 cm−1. The carbonyl group causes a strong band through C=O stretching from 1700 to 1725 cm−1 [56].
Some IR vibrations give stronger signals as the aldehydes, ketones, and other carbonyl-containing compounds. The C=O stretching vibration is the strongest signal in the IR spectrum of aldehydes and ketones; this strong signal is observed at 2383.26 to 2309.20 cm−1 in the tannins extracted by an aqueous medium. Thus, the reactions of aldehydes and ketones are essential in three types: nucleophilic addition, oxidation, and reduction. Due to the resonance of the carbonyl group, where the most important reaction by aldehydes and ketones is the nucleophilic addition reaction, which has a mechanism as follows (Figure 5):
Tannins react with the basic groups of protein in the leather matrix [58]. At first, the tannin hydroxyl groups (OH) bind to the active collagen centers, which then continue to fill the interfibrillar spaces in the leather, see Figure 6.
Tannins combine with the positively charged –NH3+ in the collagen of leather and eliminate the water, according to the reactions [13].
C O O − . P . N H 3 + + T a n n i n O − + H + ⟶ C O O H . P . N H 3 + + [ T a n n i n ] O −
C O O H . P . N H 3 + + T a n n i n O − ⟶ C O O H . P . N H 3 [ T a n n i n ] O
C O O H . P . N H 3 + + T a n n i n O ⟶ C O O H . P . N H T a n n i n + H 2 O
The binding of tannins to collagens is dependent on temperature, pH, and the molecular weight of the tannins. Therefore, it controls bacterial growth, avoids bad odors, and makes the leather stronger [14,61].
Figure 6 shows that the hydroxyl groups of the catechins present in the molecular structure of the tannins form Van der Walls electrostatic molecular interactions and hydrogen bonds with the amino and carboxylic groups present in collagen. This type of molecular interaction makes the tanning process efficient because condensed tannins have a high resistance to detanning; thus, it is suitable to tan leather in the industry [13,14].

3.4. Scanning Electron Microscopy (SEM) Analysis

The analysis by scanning electron microscopy and analysis of elemental composition by X-ray spectroscopy of the energy dispersion of the tannins in each species are reported in Figure 7.
The sample’s homogeneity was observed in the SEM image of the tannin extract of the Pinus ayacahuite species. The elemental analysis and chemical composition using X-rays determined that this tree species is presented mainly as carbon (71.61%) and oxygen (28.39%). In addition, it is possible to identify the presence of silver ions in the X-ray analysis, although this was not quantifiable.
For the Pinus dauglasiana species, the EDS analysis showed mainly carbon (61.17%) and oxygen (38.83%). For the Pinus patula species, the elemental analysis showed mainly carbon (65%), nitrogen (18.94%), oxygen (13.15%), cadmium (1.93%), and iodine (0.98%).
For the Pinus pseudostrobus species, the EDS analysis determined the presence of carbon (68.24%), oxygen (26.18%), aluminum (4.07%), and silver (1.51%). The elemental analysis of Pinus rudis using X-rays, determined the presence of carbon (68.16%), oxygen (30.88%), iodine (0.67%), and traces of silver ions. However, it was not significant.
Each atomic structure has a distinctive energy pattern, thus, generating unique electromagnetic emission spectra depending on the atoms present in the sample. In this way, energy-dispersive X-ray spectroscopy allows the characterization of the elemental composition of each of the tannins [59].
It is necessary to emphasize that plants contain a great variety of elements in various chemical forms in addition to the significant elements of their organic structure, which are carbon, hydrogen, and oxygen. Many minerals, particularly nitrogen, sulfur, and calcium, form part of its organic structure. However, other minerals may be present simply because they are absorbed as ions along with soil water and tend to accumulate as dissolved or stored ionic substances or as precipitates in tissue [60,61]. The cases of Pinus patula, Pinus pseudostrobus, and Pinus rudis present secondary ions of minerals in the analyzed tannic extracts.
As previously mentioned, SEM scanning electron microscopy is a non-destructive surface characterization technique that provides the morphological information and chemical composition of materials [62]. This modern equipment works with conductive, non-conductive, dry samples, and wet samples. However, the success of the analysis is highly dependent on sample preparation [63,64].

4. Conclusions

The content of extractable material in the bark of P. patula, P. pseudostrobus, P. ayacahuite, P. douglasiana, and P. rudis was evaluated using ethanol extraction and aqueous extraction methods. The species with the highest proportion of condensed tannins were P. patula (5.14%) and P. ayacahuite (4.71%) in the ethanolic extraction and P. patula (1.46%) and P. douglasiana (1.41%) in the aqueous extraction.
The solubility of an organic compound is directly related to its molecular structure and the polarity of the molecular bonds of the solute and solvent. Actually, the solubility of solids or liquids in another liquid will only occur if the interaction between the solute and the solvent is sufficiently high to promote the rupture of the solute–solute and solvent–solvent interactions.
However, the tannins extracted in the aqueous medium using water present a more intense signal in the FTIR analyses at 2383.26 to 2309.20 cm−1, which corresponds to a higher concentration of highly reactive carbonyl, aldehyde, and ketone groups. These can make the leather tanning process efficient and can be used in other industrial processes, such as adhesives, among others due to the high stability of the bonds.
These results allow the interest of future research to be concentrated on the condensed tannins present in these species and find applications for the large amounts of plant residue generated by forest harvesting in the community of Ixtlan de Juarez, Oaxaca, Mexico.
SEM scanning electron microscopy results provided information on the main chemical composition of these extracts. These compounds are formed of mainly carbon and oxygen, as well as hydrogen. However, some species presented mineral ions in small amounts within their composition due to the absorption of these minerals from the soil.

Author Contributions

Conceptualization, R.F.-R., F.R.-A., O.F.M.-R. and L.H.R.-T.; methodology, F.R.-A., O.F.M.-R., S.O.R.-C., R.G.-P., L.H.R.-T. and R.P.-K.; software, S.O.R.-C., F.R.-A., M.A.S.-M., M.E.S.-M. and J.P.-V.; validation, R.F.-R., F.R.-A., M.E.S.-M. and M.A.S.-M.; formal analysis, O.F.M.-R., R.G.-P., L.H.R.-T. and R.P.-K.; investigation, R.F.-R., F.R.-A., L.H.R.-T. and R.G.-P.; resources, R.F.-R., L.H.R.-T., F.R.-A. and R.G.-P.; data curation, J.P.-V., M.A.S.-M., R.G.-P. and S.O.R.-C.; writing—original draft preparation, S.O.R.-C., R.F.-R., F.R.-A. and R.G.-P.; writing—review and editing, R.F.-R., F.R.-A. and L.H.R.-T.; visualization, R.F.-R. and F.R.-A.; supervision, R.F.-R. and F.R.-A.; project administration, R.F.-R. and F.R.-A.; funding acquisition, R.F.-R., F.R.-A., L.H.R.-T., M.A.S.-M. and R.P.-K. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding. Funding was received from the resources of the University of Sierra Juarez and the National Technological Institute of Mexico–Oaxaca.

Data Availability Statement

Data is contained within the article.

Acknowledgments

The authors thank the microscopy laboratory of the Technological Institute of Oaxaca for the facilities granted to carry out the analyzes, and the community of Ixtlán de Juárez for the facilities granted for the realization of this study.

Conflicts of Interest

The authors declare no conflict of interest.

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Figure 1. Tannin classification. The numbers in the figure in the molecular structures of catechins indicate the possible order of reaction in the molecules and the importance of each benzene ring for molecular interactions to occur.
Figure 1. Tannin classification. The numbers in the figure in the molecular structures of catechins indicate the possible order of reaction in the molecules and the importance of each benzene ring for molecular interactions to occur.
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Forests 14 01433 g002 550
Figure 2. Location of the study area.
Figure 2. Location of the study area.
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Figure 3. Tannin extraction process.
Figure 3. Tannin extraction process.
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Forests 14 01433 g004a 550Forests 14 01433 g004b 550Forests 14 01433 g004c 550
Figure 4. FTIR spectra of the ethanol-extracted tannins in the bark (left) and FTIR spectra of the tannins extracted with distilled water in the bark of each species (right). The color in the figure is to differentiate the importance of each spectrum and the most important points in an FTIR analysis.
Figure 4. FTIR spectra of the ethanol-extracted tannins in the bark (left) and FTIR spectra of the tannins extracted with distilled water in the bark of each species (right). The color in the figure is to differentiate the importance of each spectrum and the most important points in an FTIR analysis.
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Figure 5. Mechanism of resonance for carbonyl group [57]. The colors in the reaction mechanism means the importance of the degree of reactivity that each chemical element has. This is very important in the language of organic chemistry.
Figure 5. Mechanism of resonance for carbonyl group [57]. The colors in the reaction mechanism means the importance of the degree of reactivity that each chemical element has. This is very important in the language of organic chemistry.
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Forests 14 01433 g006 550
Figure 6. Molecular interaction of a tannin structure with leather collagen. The colors in the molecular structures correspond to the most reactive functional groups of the catechins. The arrow indicates that the functional groups at the ends are the most reactive and react with the ketone and amine groups that are present in the molecular structure of collagen. Carbon 4 of the benzene ring 2 reacts with carbon 8 of the first benzene ring of a simple catechin structure, thus forming the molecular structure of tannins or polyphenols.
Figure 6. Molecular interaction of a tannin structure with leather collagen. The colors in the molecular structures correspond to the most reactive functional groups of the catechins. The arrow indicates that the functional groups at the ends are the most reactive and react with the ketone and amine groups that are present in the molecular structure of collagen. Carbon 4 of the benzene ring 2 reacts with carbon 8 of the first benzene ring of a simple catechin structure, thus forming the molecular structure of tannins or polyphenols.
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Figure 7. Scanning electron microscopy (SEM) micrograph of each species’ tannin extracts. The analyzed field area was 8 µm and 26.9 µm thick, with 10,000× magnification.
Figure 7. Scanning electron microscopy (SEM) micrograph of each species’ tannin extracts. The analyzed field area was 8 µm and 26.9 µm thick, with 10,000× magnification.
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Table
Table 1. Dasometric data and geographic location of the studied species.
Table 1. Dasometric data and geographic location of the studied species.
SpecieAge (Years)DBH (cm)H (m)Geographic Location
Pinus ayacahuite4133.517.517°27′38.1″ N
96°25′31.9″ O
Pinus douglasiana4535.718.4517°27′38.1″ N
96°25′31.9″ O
Pinus patula443621.617°27′38.1″ N
96°25′31.9″ O
Pinus pseudostrobus5548.518.3517°27′41.3″ N
96°25′02.6″ O
Pinus rudis523621.4517°24′15.3″ N
96°29′29.9″ O
DBH: diameter at breast height; H: height.
Table
Table 2. Average yields of ethanolic extracts.
Table 2. Average yields of ethanolic extracts.
SpeciesTotal Extract (%)Stiasny Number (%)Condensed Tannins (%)
P. ayacahuite8.44 (0.128) b55.74 (0.529) b4.71 (0.108) ab
P. douglasiana14.53 (0.176) a31.29 (2.001) d4.55 (0.346) b
P. patula7.54 (0.339) c68.14 (0.959) a5.14 (0.271) a
P. pseudostrobus4.92 (0.109) d47.12 (0.268) c2.32 (0.042) c
P. rudis7.58 (0.238) c8.61 (0.682) e0.65 (0.065) d
The mean obtained for total extract, Stiasny number, and condensed tannins for each species are presented. Values in parentheses represent the standard deviation. Mean values per column with different letters indicate significant statistical differences (p ≤ 0.05).
Table
Table 3. Average yields of aqueous extracts.
Table 3. Average yields of aqueous extracts.
SpeciesTotal Extract (%)Stiasny Number (%)Condensed Tannins (%)
P. ayacahuite2.99 (0.022) b44.92 (0.539) b1.34 (0.026) b
P. douglasiana3.19 (0.338) a44.29 (1.387) b1.41 (0.031) ab
P. patula2.26 (0.046) c64.60 (3.306) a1.46 (0.049) a
P. pseudostrobus1.16 (0.030) e33.78 (1.247) c0.35 (0.026) b
P. rudis1.70 (0.057) d8.18 (0.667) d0.14 (0.006) d
The means obtained for total extract, Stiasny number, and condensed tannins for each species. Values in parentheses represent the standard deviation. Mean values per column with different letters indicate significant statistical differences (p ≤ 0.05).
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Feria-Reyes, R.; Ramírez-Cruz, S.O.; Ruiz-Aquino, F.; Robledo-Taboada, L.H.; Sánchez-Medina, M.A.; Mijangos-Ricárdez, O.F.; Gabriel-Parra, R.; Suárez-Mota, M.E.; Puc-Kauil, R.; Porcallo-Vargas, J. Pine Bark as a Potential Source of Condensed Tannin: Analysis through Fourier Transform Infrared Spectroscopy (FTIR), Scanning Electron Microscopy (SEM), and Energy Dispersive X-ray (EDX). Forests 2023, 14, 1433. https://doi.org/10.3390/f14071433

AMA Style

Feria-Reyes R, Ramírez-Cruz SO, Ruiz-Aquino F, Robledo-Taboada LH, Sánchez-Medina MA, Mijangos-Ricárdez OF, Gabriel-Parra R, Suárez-Mota ME, Puc-Kauil R, Porcallo-Vargas J. Pine Bark as a Potential Source of Condensed Tannin: Analysis through Fourier Transform Infrared Spectroscopy (FTIR), Scanning Electron Microscopy (SEM), and Energy Dispersive X-ray (EDX). Forests. 2023; 14(7):1433. https://doi.org/10.3390/f14071433

Chicago/Turabian Style

Feria-Reyes, Rossy, Sergio Obed Ramírez-Cruz, Faustino Ruiz-Aquino, Luis Humberto Robledo-Taboada, Marco Antonio Sánchez-Medina, Oscar Francisco Mijangos-Ricárdez, Rosalío Gabriel-Parra, Mario Ernesto Suárez-Mota, Ramiro Puc-Kauil, and Jhazeel Porcallo-Vargas. 2023. "Pine Bark as a Potential Source of Condensed Tannin: Analysis through Fourier Transform Infrared Spectroscopy (FTIR), Scanning Electron Microscopy (SEM), and Energy Dispersive X-ray (EDX)" Forests 14, no. 7: 1433. https://doi.org/10.3390/f14071433

APA Style

Feria-Reyes, R., Ramírez-Cruz, S. O., Ruiz-Aquino, F., Robledo-Taboada, L. H., Sánchez-Medina, M. A., Mijangos-Ricárdez, O. F., Gabriel-Parra, R., Suárez-Mota, M. E., Puc-Kauil, R., & Porcallo-Vargas, J. (2023). Pine Bark as a Potential Source of Condensed Tannin: Analysis through Fourier Transform Infrared Spectroscopy (FTIR), Scanning Electron Microscopy (SEM), and Energy Dispersive X-ray (EDX). Forests, 14(7), 1433. https://doi.org/10.3390/f14071433

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